Rem uncouples excitation–contraction coupling in adult skeletal muscle fibers

Excitation–contraction (EC) coupling is the physiological event in which muscle converts an electrical signal (plasma membrane depolarization) into mechanical work (muscle contraction). In the case of skeletal muscle, depolarization-induced conformational rearrangements within the L-type Ca²⁺ channel complex (Ca_V1.1) are coupled to gating of the type 1 ryanodine-sensitive Ca²⁺ release channel (RYR1) of the SR (Schneider and Chandler, 1973; Ríos and Brum, 1987; Tanabe et al., 1988). The resultant Ca²⁺ efflux from the SR into the myoplasm via RYR1 activates the contractile filaments. Because SR Ca²⁺ release occurs rapidly in response to depolarization and independently of transient Ca²⁺ fluctuations, a conformational coupling mechanism appears to support communication between the two channels (see Bannister and Beam, 2013).

Although the roles of Ca_V1.1 and RYR1 as voltage sensor and SR Ca²⁺ release channel, respectively, have been established for quite some time (Tanabe et al., 1988; Nakai et al., 1996), the molecular mechanics that support conformational coupling between these two channels remain undefined. One candidate structure to mediate such coupling is the intracellular segment that links repeats II and III of the principal α_1S subunit of Ca_V1.1 (Tanabe et al., 1990; Lu et al., 1994; Nakai et al., 1998; Wilkens et al., 2001). Another viable candidate is the auxiliary β_1a subunit of the Ca_V1.1 heteromultimer. In this regard, β_1a is firmly established as being essential for EC coupling, as genetic deletion of β₁ abolishes voltage-dependent SR Ca²⁺ release in both mammals and bony fish (Gregg et al., 1996; Ono et al., 2004; Schredelseker et al., 2005, 2009). Moreover, purified β_1a subunits and β_1a peptide fragments bind RYR1 in vitro and/or activate RYR1 in planar lipid bilayers (Cheng et al., 2005; Rebbeck et al., 2011; Karunasekara et al., 2012; Hernández-Ochoa et al., 2014). Still, the key roles of β_1a in trafficking Ca_V1.1 to the plasma membrane (Gregg et al., 1996; Strube et al., 1996) and in the ultrastructural organization of Ca_V1.1 into the tetradic arrays prerequisite for EC coupling (Schredelseker et al., 2005, 2009; Dayal et al., 2010, 2013; Eltit et al., 2014) have made testing a direct role for β_1a in communication between the voltage-sensing components of Ca_V1.1 and RYR1-mediated SR Ca²⁺ release highly problematic.

Members of the RGK (Rad, Rem, Rem2, Gem/Kir) family of monomeric G proteins inhibit L-type Ca²⁺ channels in a variety of physiological systems via interactions that occur primarily with the β subunit (Béguin et al., 2001, 2007; Finlin et al., 2003, 2006; Murata et al., 2004; Bannister et al., 2008; Yang et al., 2012; Romberg et al., 2014; Xu et al., 2015; reviewed recently by Yang and Colecraft, 2013). In the present study, we have examined the impact of Rem on EC coupling in adult mouse flexor digitorum brevis (FDB) fibers overexpressing Rem via in vivo electroporation (DiFranco et al., 2007). Using this approach, we have found that Rem effectively uncouples the Ca_V1.1 voltage sensor from RYR1-mediated SR Ca²⁺ release through its interaction with β_1a. Specifically, Rem markedly reduces voltage-induced myoplasmic Ca²⁺ transients without appreciable effects on Ca_V1.1 targeting, intramembrane charge movement, or SR Ca²⁺ store content.

A cDNA encoding a CFP-rabbit Ca_V1.1 α_1S-subunit (GenBank accession no. X05921) fusion construct was created by swapping out YFP for CFP in an existing YFP–α_1S fusion construct (Papadopoulos et al., 2004). The cDNA segment encoding CFP was excised from the parent pECFP-C1 vector (Takara Bio Inc.) using NheI and HindIII (761 bp). Likewise, YFP was removed from the YFP–α_1S fusion construct using the same restriction enzymes, linearizing the pEYFP-C1 backbone and the α_1S-coding sequence (9,555 bp). The CFP-encoding segment was then re-ligated into the linearized vector carrying the α_1S-coding sequence (final, 10,316 bp).

The constructions of CFP–rabbit β_1a and YFP–rabbit β_1a (both GenBank accession no. M25514) were described previously by Leuranguer et al. (2006); CFP-β_1a, YFP-β_1a, and CFP-Ca_V1.1 were all provided by K.G. Beam (University of Colorado Denver-Anschutz Medical Campus, Aurora, CO).

The construction of V-Rem AAA (RefSeq accession no. NP_033073) was described previously by Beqollari et al. (2015). Restriction digests and sequencing were used to verify all constructs.

All procedures involving mice were approved by the University of Colorado Denver-Anschutz Medical Campus Institutional Animal Care and Use Committee. cDNA plasmids encoding YFP, CFP-α_1S, CFP-β_1a, V-Rem, and/or V-Rem AAA were delivered to FDB fibers of anesthetized 2–3-mo-old male C57BL/6J mice (The Jackson Laboratory) via an in vivo electroporation protocol similar to that originally described by DiFranco et al. (2007). In brief, 10 µl of 2 mg/ml hyaluronidase solution was injected into the FDB muscle with a 30-gauge hypodermic needle. After 1 h, mice were re-anesthetized and 20 µl cDNA (3–5 µg/µl) was injected into the muscle. 5 min later, two gold-plated acupuncture needle electrodes (Lhasa OMS) coupled to an isolated pulse stimulator (A-M Systems) were placed subcutaneously near the proximal and distal tendons of the muscle (∼1 cm apart). cDNAs were then electroporated into the FDB muscle with 20 100-V, 20-ms pulses delivered at 1 Hz. For assessment of SR Ca²⁺ stores, the transfection mixture also contained 5 µg pmCherry-C1 (Takara Bio Inc.) as a means to identify successfully transfected fibers after loading with Fluo 3-AM dye (Invitrogen; see below).

Electroporated (9–10 d after transfection) FDB muscles were dissected in cold rodent Ringer’s solution (mM: 146 NaCl, 5 KCl, 2 CaCl₂, 1 MgCl₂, and 10 HEPES, pH 7.4 with NaOH). Muscles were then digested in a collagenase solution (mM: 155 Cs-aspartate, 10 HEPES, and 5 MgCl₂, pH 7.4 with CsOH, supplemented with 1 mg/ml BSA [Sigma-Aldrich] and 1 mg/ml collagenase type IA [Sigma-Aldrich]) with agitation at 37°C for ∼1 h. Immediately after digestion, the collagenase solution was replaced with a dissociation solution (mM: 140 Cs-aspartate, 10 Cs₂EGTA, 10 HEPES, and 5 MgCl₂, pH 7.4 with CsOH, supplemented with 1 mg/ml BSA), and muscles were triturated gently with a series of fire-polished glass pipettes of descending bore. Dissociated FDB fibers destined for whole-cell patch-clamp experiments were then plated onto ECL (EMD Millipore)-coated 35-mm plastic culture dishes (Falcon). For imaging, fibers were allowed to settle onto laminin (Invitrogen)-coated 35-mm culture dishes with glass coverslip bottoms (MatTek). Experiments were performed with FDB fibers 1–6 h after dissociation; successfully transfected fibers were identified by the presence of YFP or Venus fluorescence.

Patch pipettes were fabricated from borosilicate glass and had resistances of ≤1.0 MΩ when filled with internal solution, which consisted of (mM): 140 Cs-aspartate, 10 Cs₂-EGTA, 5 MgCl₂, and 10 HEPES, pH 7.4 with CsOH; fibers were dialyzed in the whole-cell configuration for >20 min before recording. For recording of L-type Ca²⁺ currents, the external solution contained (mM): 145 TEA-methanesulfonic acid, 10 CaCl₂, 10 HEPES, 2 MgSO₄, 1 4-aminopyridine, 0.1 anthracene-9-carboxylic acid, and 0.002 tetrodotoxin, pH 7.4 with TEA-OH. For measurement of charge movements, the bath contained (mM): 145 TEA-methanesulfonic acid, 10 CaCl₂, 10 HEPES, 2 MgSO₄, 1 4-aminopyridine, 0.1 anthracene-9-carboxylic acid, 0.002 tetrodotoxin, 1 LaCl₃, and 0.5 CdCl₂, pH 7.4 with TEA-OH. Linear components of leak and capacitive current were corrected with −P/4 online subtraction protocols. Output filtering was at 2–5 kHz, and digitization was either at 5 kHz (currents) or 10 kHz (charge movements). Cell capacitance was determined by integration of a transient from −80 to −70 mV using Clampex 10.3 (Molecular Devices) and was used to normalize charge movement (nC/µF) and current amplitude (pA/pF). The average value of C_m was 2.3 ± 0.1 nF (n = 48 fibers). To minimize voltage error, the time constant for decay of the whole-cell capacity transient (τ_m) was reduced as much as possible using the analogue compensation circuit of the amplifier; the average values of τ_m and R_a were 1.0 ± 0.02 ms and 467 ± 26 kΩ, respectively. Q_ON was then normalized to C_m and plotted as a function of test potential (V), and the resultant Q-V relationship was fitted according to:

where Q_max is the maximal Q_ON, V_Q is the potential causing movement of half the maximal charge, and k_Q is a slope parameter. Peak currents were normalized to C_m, and the resultant I-V was fitted according to:

where I is the normalized current for the test potential V, V_rev is the reversal potential, G_max is the maximum Ca²⁺ channel conductance, V_1/2 is the half-maximal activation potential, and k_G is the slope factor. All electrophysiological and Ca²⁺-imaging experiments were performed at room temperature (∼25°C).

Voltage-induced changes in myoplasmic Ca²⁺ were recorded from FDB fibers with Fluo 3 single-wavelength Ca²⁺ indicator dye (Invitrogen). The pentapotassium salt form of the dye was added to the standard internal solution (see above) for a final concentration of 200 µM. The external solution contained (mM): 145 TEA-methanesulfonic acid, 10 CaCl₂, 10 HEPES, 2 MgSO₄, 1 4-aminopyridine, 0.1 anthracene-9-carboxylic acid, and 0.002 tetrodotoxin, pH 7.4 with TEA-OH. After entry into the whole-cell configuration, a waiting period of no less than 20 min was used to allow the dye to diffuse into the cell interior. A 100-W mercury illuminator and a set of fluorescein filters were used to excite the dye present in the voltage-clamped fiber. A computer-controlled shutter was used to block illumination in the intervals between test pulses. Fluorescence emission was measured by means of a fluorometer (Biomedical Instrumentation Group, University of Pennsylvania). Fluorescence data are expressed as ΔF/F, where ΔF represents the change in peak fluorescence from baseline during the test pulse, and F is the fluorescence immediately before the test pulse minus the average background fluorescence. The peak value of the fluorescence change (ΔF/F) for each test potential (V) was fitted according to:

where (ΔF/F)_max is the maximal fluorescence change, V_F is the potential causing half the maximal change in fluorescence, and k_F is a slope parameter. Only cells with transients that could be fit with Eq. 3 were used for analysis.

Dissociated FDB fibers were examined in rodent Ringer’s solution using a confocal laser-scanning microscope (LSM 510 META; Carl Zeiss). A Plan-Apochromat 63× oil-immersion objective (1.4 NA) was used to view the fiber of interest. CFP and Venus were excited with separate sweeps of the 458- and 514-nm lines, respectively, of an argon laser (30-milliwatt maximum output, operated at 50% or 6.3 A) directed to the cell via a 458/514-nm dual dichroic mirror. The emitted fluorescence was split via a 515-nm long-pass filter; CFP was directed to a photomultiplier equipped with a 465–495-nm band-pass filter, and Venus was directed to a photomultiplier equipped with a 530-nm long-pass filter. The chosen settings precluded recording of fluorescence bleed between CFP and Venus, because CFP is not excited at 514 nm and Venus emission is negligible between 465 and 495 nm (see Papadopoulos et al., 2004). Confocal fluorescence intensity data were recorded as the average of eight line scans per pixel and digitized at 8 bits, with photomultiplier gain adjusted such that maximum pixel intensities were no more than ∼70% saturated.

FDB fibers were loaded with 5 µM Fluo 3-AM and 0.05% pluronic acid (both from Invitrogen) dissolved in rodent Ringer’s solution for 35 min at 37°C. Fibers were then washed three times in rodent Ringer’s solution with gentle agitation. After a 10-min de-esterification period, Fluo 3-AM–loaded cells bathed in rodent Ringer’s solution were placed on the stage of the LSM 510 META microscope and viewed with a 10× 0.3-NA objective (Carl Zeiss). Fluo 3-AM was excited with the 488-nm line of an argon laser (30-milliwatt maximum output, operated at 50% or 6.3 A, attenuated to 5%). The emitted fluorescence was directed through a dual 488/543 dichroic mirror to a photomultiplier equipped with a 500–530-nm band-pass filter. SR Ca²⁺ release was induced by 1 mM 4-chloro-m-cresol (4-CmC; Pfaltz & Bauer) delivered via a manually operated, gravity-driven global perfusion system. Fluorescence amplitude data are expressed as ΔF/F, where F represents the baseline fluorescence before application of 4-CmC, and ΔF represents the change in peak fluorescence during the application of 4-CmC.

Low (<20) passage tsA201 cells were propagated in culture medium containing 90% DMEM (Thermo Fisher Scientific), 10% defined fetal bovine serum (GE Healthcare), and 100 µg/ml penicillin-streptomycin (Life Technologies). Cells were trypsinized twice weekly and replated onto 35-mm culture dishes at ∼20% confluence. Lipofectamine 2000 (Life Technologies) was used to transfect these cells within 3–5 d of plating. The transfection mixture contained expression plasmids encoding rat Ca_V1.3, rabbit β_1a, and rat α₂δ1 channel subunits at 1 µg of each cDNA per dish. The transfection mixture also contained a plasmid-encoding Venus–Rem construct (1 µg/dish; see above) or YFP (30 ng/dish; Takara Bio Inc.). The day after transfection, cells were trypsinized and replated onto 35-mm plastic for experiments the next day.

tsA201 cells expressing YFP-β_1a, YFP-β_1a/V-Rem, or YFP-β_1a/V-Rem AAA were lysed into 300 µl of lysis buffer (mM: 50 Tris-HCl, pH 7.5, 100 NaCl, 1 MgCl₂, 1 DTT, and 0.2% Tween-20) supplemented with 0.1 mM iodoacetamide and 1 mM phenylmethylsulfonyl fluoride (both from Thermo Fisher Scientific). After insoluble material was removed by centrifugation, the homogenates were incubated with a monoclonal antibody directed to Rem (1:200; Santa Cruz Biotechnology, Inc.) for 4–6 h with gentle agitation followed by an overnight incubation with protein A agarose beads (Santa Cruz Biotechnology, Inc.). The agarose beads were then washed twice with lysis buffer and collected after gentle centrifugation at 2,500 rpm. The beads were then resuspended in 30 µl of 1% SDS buffer (Bio-Rad Laboratories) and subjected to SDS-PAGE analysis. Proteins were transferred into a nitrocellulose membrane, blocked with 3% nonfat dry milk (Kroger) in PBS-Tween, and incubated overnight at 4°C with monoclonal antibodies directed to either X(G)FP (1:1,500; Antibodies Inc.) or Rem (1:500). The nitrocellulose membrane was then washed three times with PBS-Tween and incubated at room temperature for 1 h with horseradish peroxidase–conjugated goat anti–mouse IgG (1:10,000; SouthernBiotech). Protein bands were visualized with the SuperSignal West Femto kit (Thermo Fisher Scientific) and viewed on a FluorChem HD2 scanner (Alpha Innotech). Blots were stripped using Restore Western Blot Stripping Buffer (Thermo Fisher Scientific).

Borosilicate pipettes (2.0–3.0 MΩ) were filled with internal solution, which consisted of (mM): 140 Cs-aspartate, 10 Cs₂-EGTA, 5 MgCl₂, and 10 HEPES, pH 7.4 with CsOH. The bath solution contained (mM): 145 NaCl, 10 CaCl₂, and 10 HEPES, pH 7.4 with NaOH. Electronic compensation was used to reduce the effective series resistance, and linear components of leak and capacitive current were corrected with −P/4 online subtraction protocol. Filtering and digitation were at 2 and 5 kHz, respectively. For tsA201 cell experiments, the average values of C_m, τ_m, and R_a were 20.0 ± 1.4 pF, 202.0 ± 18.9 µs, and 10.9 ± 0.9 MΩ, respectively (n = 28 cells).

All data are presented as mean ± SEM. Statistical comparisons were made by unpaired t test or by one-way ANOVA (where appropriate), with P < 0.05 considered significant. Figures were made using the software program SigmaPlot (version 11.0; SSPS Inc.).

Fig. S1 shows, qualitatively, the successful expression of both V-Rem and V-Rem AAA in FDB fibers by in vivo electroporation. Confocal fluorescence images of six different live, intact FDB fibers overexpressing V-Rem or V-Rem AAA are shown with average intensity profiles for the indicated regions of interest.

Recently, we described the effects of the RGK proteins Rad and Rem on L-type Ca²⁺ currents and intramembrane charge movement in adult FDB muscle fibers (Beqollari et al., 2014). Although both Rad and Rem inhibited L-type currents by ∼60 and ∼45%, respectively, charge movement was only reduced in fibers transfected with Rad; charge movement for Rem-expressing fibers was virtually identical to charge movement observed in naive fibers. To confirm the latter observation, we used in vivo electroporation (DiFranco et al., 2007) to transfect FDB muscles of otherwise normal 2–3-mo-old C57BL/6J mice with either YFP or a Venus-fused wild-type mouse Rem construct (V-Rem). As expected, FDB fibers overexpressing V-Rem again displayed maximal charge movement virtually identical to YFP-expressing fibers in both amplitude and voltage dependence (Fig. 1, A–C and Table 1). Both Q-V relationships were similar to that reported by Prosser et al. (2009) when La³⁺ was included in the extracellular recording solution.

Because skeletal muscle EC coupling is coupled directly to translocation of Ca_V1.l’s voltage-sensing structures (Schneider and Chandler, 1973; Ríos and Brum, 1987; García et al., 1994; Tanabe et al., 1988), we next investigated the impact of Rem on EC coupling by recording myoplasmic Ca²⁺ transients in response to membrane depolarization (as in Wang et al., 1999; Wu et al., 2012). Ca²⁺ transients recorded from fibers transfected with V-Rem were substantially reduced compared with the transients of YFP-expressing fibers (0.6 ± 0.1 ΔF/F, n = 8 vs. 1.6 ± 0.4 ΔF/F, n = 6, respectively; P < 0.001; Fig. 1, D–F). No significant effect on the voltage dependence of SR Ca²⁺ release was observed between the two groups (P > 0.05; Table 1).

As a means to determine whether the V-Rem–mediated reduction in voltage-induced Ca²⁺ release was a consequence of an altered SR Ca²⁺ store, we exposed intact fibers loaded with Fluo-3 AM dye to the RYR agonist 4-CmC. In these experiments, 1 mM 4-CmC elicited SR Ca²⁺ release that was nearly indistinguishable between FDB fibers overexpressing V-Rem and fibers expressing YFP only (5.7 ± 1.0 ΔF/F, n = 9 vs. 6.8 ± 1.8 ΔF/F, n = 5, respectively; P > 0.05; Fig. 2, A–C). The equivalent responses of YFP- and V-Rem–expressing fibers to 4-CmC suggest that depletion of SR Ca²⁺ store is an unlikely explanation for the ∼65% reduction in Ca²⁺ transient amplitude observed in V-Rem–expressing fibers.

Because Rem has been reported to alter high voltage-activated Ca²⁺ channel trafficking in heterologous systems (Béguin et al., 2007; Mahalakshmi et al., 2007; Flynn and Zamponi, 2010; Yang et al., 2010) and in cardiac myocytes (Jhun et al., 2012), one possible explanation for the disruption of EC coupling by V-Rem (Fig. 1, D and F) is that the small G protein redirects Ca_V1.1 away from triad junctions. For this reason, we examined the subcellular distribution of Ca_V1.1 α_1S and β_1a subunits in the absence and presence of coexpressed V-Rem. When expressed in FDB fibers, CFP-tagged α_1S subunits of Ca_V1.1 were targeted to transverse tubules as shown previously for YFP-tagged α_1S subunits (DiFranco et al., 2011; Fig. 3 A). The tubular distribution of CFP-α_1S was unaffected by coexpression of V-Rem (Fig. 3 B). Likewise, coexpression of V-Rem had little, if any, effect on the subcellular distribution of CFP-β_1a (Fig. 3, C and D). Interestingly, the V-Rem fluorescence extended from the transverse tubules into the region of the I band. In this regard, the subcellular distribution of V-Rem overlapped, but did not completely match, the transverse tubular distributions of Ca_V1.1 α_1S and β_1a subunits. We do not consider the presence of Rem in the I band to be an artifact of overexpression, as the related RGK protein Rad clearly targets to transverse tubules when expressed in FDB fibers via electroporation (see Beqollari et al., 2014). Moreover, this observation does not affect our interpretation of the data shown in Fig. 3: coexpression of V-Rem did not alter targeting of the channel subunits to the transverse tubules.

So far, our data indicate that Rem uncouples the Ca_V1.1 voltage sensor from RYR1-mediated SR Ca²⁺ release. However, it is unclear whether this effect of Rem is dependent on the ability of the small GTP-binding protein to interact with the β_1a subunit of the Ca_V1.1 channel complex. In this regard, three highly conserved residues of Rem (R200, L227, or H229) have been identified as being critical for interactions with the β₃-subunit isoform (Béguin et al., 2007; Puhl et al., 2014); conversion of any one of these residues to alanine severely impairs binding to β₃-subunit isoforms in both yeast-2-hybrid and coimmunoprecipitation assays. To specifically test whether Rem binds to the β_1a-subunit isoform, we engineered a V-Rem–based construct with alanines introduced at positions R200, L227, and H229 (V-Rem AAA) and compared its ability to coimmunoprecipitate with a YFP-fused β_1a construct (YFP-β_1a). In these experiments, a commercially available monoclonal Rem antibody failed to immunoprecipitate YFP-β_1a in lysates obtained from tsA201 cells transfected with only YFP-β_1a (shown in duplicate in Fig. 4 A, lanes 2 and 6). In contrast, the antibody efficiently immunoprecipitated YFP-β_1a subunits when V-Rem was coexpressed with YFP-β_1a (Fig. 4 A, lanes 3 and 7). Consistent with the earlier report of Béguin et al. (2007) showing the disruption of the Rem-β₃ interaction with alanine single-point mutants, an interaction between V-Rem AAA and β_1a was not detectable (Fig. 4 A, lanes 4 and 8). In control experiments, the Rem antibody detected similar levels of immunoprecipitated V-Rem and V-Rem AAA (Fig. 4 B, lanes 3–4 and 7–8). Comparable expression levels for YFP-β_1a, V-Rem, and V-Rem AAA mutant were confirmed in a Western blot from total lysates collected before coimmunoprecipitation (Fig. 4 C).

We next determined the functional consequences of the disruption of the Rem-β_1a interaction. In these experiments, we coexpressed YFP, V-Rem, or V-Rem AAA with Ca_V1.3 α_1D, β_1a, and α₂δ-1 subunits to detect interactions that occur within a functional L-type channel complex (we used Ca_V1.3 as a surrogate for Ca_V1.1 because of its highly efficient and consistent membrane expression in tsA201 cells; see Meza et al., 2013). Predictably, tsA201 cells expressing Ca_V1.3, β_1a, α₂δ-1, and V-Rem displayed virtually no L-type current (−3.8 ± 0.7 pA/pF at 0 mV; n = 4; Fig. 5 A). In contrast, cells expressing Ca_V1.3, β_1a, α₂δ-1, and V-Rem AAA had L-type currents nearly identical in amplitude (−78.3 ± 16.0 pA/pF, n = 15; Fig. 5, B and D) to control cells expressing the same channel subunits with a YFP transfection marker (−71.0 ± 20.0 pA/pF, n = 9; P > 0.05; Fig. 5, C and D). Successful expression of V-Rem and V-Rem AAA in tsA201 cells was indicated by Venus fluorescence (Fig. 5 E).

To establish β_1a as the mechanistic target of Rem in our experimental system, we overexpressed V-Rem AAA in FDB fibers and assayed its effects on L-type Ca²⁺ currents, intramembrane charge movement, and EC coupling. Successful expression of V-Rem AAA in FDB fibers was confirmed by Venus fluorescence, which was comparable to the fluorescence generated by V-Rem (see examples in Fig. S1). FDB fibers expressing V-Rem AAA produced sizable L-type currents that were not different than those observed in fibers expressing YFP (−9.8 ± 1.2 pA/pF, n = 6 and −9.0 ± 0.5 pA/pF, n = 5, respectively, at 20 mV; P > 0.05; Fig. 6 A). Likewise, V-Rem AAA had no obvious effect on the magnitude of gating charge movement (24.6 ± 3.4 nC/µF, n = 5; P > 0.05, ANOVA; Fig. 6 B and Table 1), although these fibers did present a steeper Q-V relationship when compared head-to-head with YFP-expressing fibers (P < 0.05, unpaired t test). Most importantly, V-Rem AAA also failed to significantly dampen SR Ca²⁺ release in response to membrane depolarization (1.4 ± 0.2 ΔF/F, n = 6; P > 0.05, ANOVA; Fig. 6 C and Table 1). Taken with the results in Figs. 4 and 5 showing that V-Rem AAA is unable to interact with β_1a, these data indicate that the near ablation of EC coupling by V-Rem (Fig. 1) is largely dependent on structural elements that are important for contact(s) with β_1a.

In this study, we found that the RGK family small G protein Rem profoundly inhibits skeletal muscle EC coupling in adult mouse FDB muscle fibers (Fig. 1, D–F). Because the observed reduction in voltage-induced SR Ca²⁺ release was not likely a consequence of altered Ca_V1.1 targeting (Fig. 3), impaired voltage sensing (Fig. 1, A–C) or a greatly depleted SR Ca²⁺ store (Fig. 2), a “communication breakdown” must have occurred between Ca_V1.1 and RYR1. An intuitive candidate locus for such EC uncoupling is the auxiliary β_1a subunit of the Ca_V1.1 heteromultimer because RGK proteins are established β-subunit–interacting partners (Béguin et al., 2001, 2007; Finlin et al., 2003, 2006; Yang and Colecraft, 2013; Puhl et al., 2014; Xu et al., 2015). Earlier work by Colecraft and colleagues has established that Rem can inhibit L-type Ca_V1.2 channels expressed in HEK 293 cells without affecting intramembrane charge movement (Yang et al., 2007, 2010), and that this particular mode of Rem-mediated inhibition is dependent solely on an interaction with the β subunit (Yang et al., 2012; Yang and Colecraft, 2013). Because Rem exclusively uses this “low P_o” gating mode to inhibit Ca_V1.1 channel function in differentiated muscle fibers (Beqollari et al., 2014), the observed impairment of EC coupling by Rem is almost certainly dependent on a Rem-β_1a interaction. The inability of V-Rem AAA, a Rem construct lacking key structural elements for β binding and channel inhibition (Figs. 4 and 5, respectively), to reduce EC coupling provides additional support for this assertion (Fig. 6).

In addition to inhibiting EC coupling, V-Rem also reduced L-type current in FDB fibers (Fig. 6 A; Beqollari et al., 2014). Because L-type Ca²⁺ entry has been found to contribute to SR Ca²⁺ store refilling in myotubes (Cherednichenko et al., 2004) and in differentiated muscle fibers (Lee et al., 2015), it is not beyond possibility that SR stores may be partially depleted in V-Rem–expressing fibers. However, the nearly equivalent responses of YFP- and V-Rem–expressing fibers to 4-CmC (Fig. 2) support the idea that such a mechanism is unlikely to account for the observed effect of Rem on voltage-induced SR Ca²⁺ release. Likewise, an acute contribution from L-type Ca²⁺ flux via the channel is also improbable, as the ΔF/F-V curves for YFP- and Rem AAA–expressing fibers both displayed sigmoidal dependencies on voltage, a hallmark indication of skeletal-type EC coupling (see Fig. 6 C, right). If Ca²⁺ flux were making a small contribution to the transients, its loss could not likely explain the nearly 65% decrease in SR Ca²⁺ release resulting from coexpression of V-Rem.

Strong circumstantial, but by no means definitive, evidence exists supporting the hypothesis that β_1a is directly involved in Ca_V1.1–RYR1 communication (see Rebbeck et al., 2014). In particular, expression of β_1a is essential for EC coupling and enhances L-type current amplitude considerably (Gregg et al., 1996; Strube et al., 1996). Unfortunately, these early results obtained with myotubes cultured from β₁ null mice have been difficult to interpret because membrane expression of the principal α_1S subunit of Ca_V1.1 was severely compromised. The confounding obstacle of poor α_1S trafficking in β₁ null mice was overcome by elegant work with the effectively β₁ null relaxed zebrafish mutant line. In the relaxed system, unpartnered α_1S subunits trafficked somewhat more effectively to plasma membrane–SR junctions than in mice (Schredelseker et al., 2005). The improved membrane expression of Ca_V1.1 enabled meticulous ultrastructural examination of relaxed junctions, revealing that β_1a is required to organize Ca_V1.1 into the tetrad arrays that are prerequisite for EC coupling.

Beyond ultrastructure, the zebrafish model system poses nearly the same challenges to deciphering the function of β_1a as does the β₁ null mouse model. Specifically, the introduction of chimeric β_1a constructs or other Ca_Vβ isoforms has been highly useful in the identification of functionally important domains, but information regarding essential intermolecular interactions remains frustratingly difficult to glean (Beam and Bannister, 2010). In efforts to avoid such ambiguity, in vitro approaches have been used to identify interactions of potential functional significance between β_1a and RYR1. Indeed, purified full-length β_1a subunits do bind fragments of RYR1 in vitro (Cheng et al., 2005; Rebbeck et al., 2011), and a peptide corresponding to β_1a residues V490–M524 increases RYR1 P_o when applied to lipid layers (Karunasekara et al., 2012). Likewise, dialysis of FDB fibers with a slightly shorter peptide (V490–M508) peptide potentiates both EC coupling and L-type Ca²⁺ current by nearly 50% (Hernández-Ochoa et al., 2014). Although the use of β_1a-based peptide approaches has provided support for the idea that β_1a residues V490–M508 are involved in transmitting the signal between Ca_V1.1 and RYR1, the interpretation of these results has been somewhat limited because of uncertainty of substrate and lack of peptide specificity; one must take into account that a variety of small peptides binds to the ginormous ∼2.3-MDa RYR1 tetramer and/or modulates RYR1 P_o in lipid bilayers (e.g., peptides corresponding to the A domain of the Ca_V1.1 II–III linker, Imperatoxin A, Maurocalcine; El-Hayek and Ikemoto, 1998; Gurrola et al., 1999; Fajloun et al., 2000; Nabhani et al., 2002; Chen et al., 2003; Cui et al., 2009).

In light of the frustrating limitations of the experimental approaches described above, new strategies are needed to further investigate the role of β_1a in skeletal-type EC coupling. The use of wild-type Rem or modified Rem constructs to probe junctional architecture represents such an advance because the small G protein disrupts Ca_V1.1–RYR1 communication in intact, differentiated muscle fibers without deleting or altering the peptide sequences of the endogenous components of the Ca²⁺ release unit (CRU). Obviously, the next step in this line of investigation is to determine the precise mechanism by which Rem cuts communication between Ca_V1.1 and RYR1. Based on what is currently known, β_1a coordinates the juxtaposition of Ca_V1.1 with RYR1 in tetrads (Fig. 7 A). So, it is quite possible that the Rem–β_1a interaction merely impairs the ability of β_1a to facilitate the ultrastructural configuration of Ca_V1.1 and RYR1 that is requisite for conformational coupling (Fig. 7 B). However, the preservation of tetrad arrays in fibers overexpressing Rem would indicate that the RGK protein is exerting its inhibitory influence on β_1a within the intact CRU, which in turn would imply that conformational changes in β_1a are involved in Ca_V1.1–RYR1 coupling (Fig. 7 C). A correlate of the latter interpretation would be that other structures (e.g., II–III loop of the α_1S subunit) thought to be involved in transmission of the EC coupling signal are adversely impacted by Rem-induced conformational changes in β_1a. Of course, these ideas remain to be tested. In this regard, our current observations provide a new means for the investigation of the β_1a subunit as mediator of the communication between Ca_V1.1 and RYR1 that underlies EC coupling skeletal muscle.

We thank Mr. M.P. Scheele for technical assistance, Drs. S.R. Ikeda and H.L. Puhl, III, for sharing the Venus-Rem expression plasmid, and Drs. K.G. Beam and D. Oskar for continued support.

This work was supported by grants from the National Institutes of Health (AG038778 to R.A. Bannister), the Colorado Clinical and Translational Sciences Institute (J-11-122 to R.A. Bannister), the Boettcher Foundation (to R.A. Bannister), and CONACyT (169006 to U. Meza). D. Beqollari received a stipend from 2T32AG000279-11 (to R.S. Schwartz, University of Colorado Denver-AMC Department of Medicine-Geriatrics Division). Confocal images were acquired in the University of Colorado Denver-AMC Advanced Light Microscopy Core (funded in part by National Institutes of Health/NCRR Colorado CTSI grant UL1 RR025780).

The authors declare no competing financial interests.

Author contributions: D. Beqollari and R.A. Bannister designed research, performed research, analyzed data, and wrote the paper. C.F. Romberg and D. Filipova performed experiments and analyzed data. U. Meza and S. Papadopoulos analyzed data and wrote the paper.

Richard L. Moss served as editor.